Aim 1 Dozens of proteins control the docking and fusion of exocytic vesicles in neurons and endocrine cells. The identity and roles of many of these proteins have been assigned through a combination of genetics, biochemistry, and electrophysiology. However, the spatial organization, heterogeneity, and dynamics of these proteins have yet to be determined. Finding this organization is key to understanding how proteins regulate these systems in healthy cells and might malfunction in disease. Thus, we aim to map key proteins proposed to act during exocytosis in endocrine cells. To accomplish this, we developed a combination of high-throughput live cell imaging, super-resolution fluorescence imaging, and electron microscopy. Through this multi-modal imaging approach, the location, dynamics, and occupancy of individual proteins were mapped at specific populations of vesicles in cells and compared to the underlying cellular architecture that organizes the plasma membrane. This allowed us to determine the fundamental organization of the exocytic membrane system and how specific molecular components responsible for vesicle trafficking and fusion assemble together and function. Specifically, we have been using two-color TIRF microscopy and high-throughput image analysis to detect and characterize over 80 different proteins that have potential roles at exocytic sites. Through this work we developed a universal map of the proteins that control exocytosis and provide a new global network level analysis of vesicle fusion. We were able to identify unique classes of key regulatory molecules that strongly associate with the vast majority of docked exocytic vesicles in endocrine chromaffin and beta cells. Key proteins we identified were Rabs and Rab effectors, SNARE proteins, and several SNARE modulators. The above mapping work was done on single images. Thus, to determine the cellular dynamics of these components we imaged the live-cell local changes of proteins at the exact moment of fusion. In these microscopy studies we discovered an unexpected rapid recruitment of several important endocytic proteins and lipids to sites of exocytosis. These molecules include the regulatory lipid PIP2, and the proteins dynamin, amphiphysin, syndapin, and endophilin. We further discovered that mutations to several of these proteins altered the kinetics of cargo release from single insulin-containing dense-core vesicles in cultured beta cells. Our hypothesis is that these proteins (dynamin, syndapin, amphiphysin, and endophilin) regulate the dilation of the fusion pore to control the amount of cargo released during single exocytic fusion events in cells of the pancreatic islets. Along with these studies we have developed a new super-resolution correlative light and electron microscopy method (CLEM). This CLEM method allows us to image the location of identified proteins in the nano-scale structural context of the cellular environment. Specifically, we have succeeded in developing a robust pipeline for imaging the plasma membrane of human cells with both fluorescence microscopy (TIRF) and transmission electron microscopy (TEM) of platinum replicas. These studies are allowing us to build structural models for how single organelles are organized and how these complexes regulate exocytosis, a central process for commination among cells and tissues in the body. We have expanded our research in this aim to the study of the large terminals of primary bipolar neurons from the retina. These terminals can be over 10 microns in diameter which make them ideal for studying the structure of the synaptic space. In these studies we discovered a new synaptic cytoskeletal structure. We found a thick marginal band of microtubules that extends down from the axon and loops throughout the synaptic terminal. This microtubule loop is structurally similar to those found in red blood cells. Using 3D confocal fluorescence, block-face serial scanning electron microscopy, and transmission electron microscopy we fully characterized this structure. We found that this microtubule structure associates with a substantial population of mitochondria in the terminals. Drugs that inhibit microtubule-based kinesin motors stalled mitochondria in the axon. From this work we conclude that this prominent microtubule band is important for the transport and localization of critical mitochondria into the presynaptic space to provide the sustained energy necessary for continuous transmitter release in these giant synaptic terminals. Aim 2 We discovered that in endocrine cells vesicle material is captured on a dense network of pre-formed clathrin-coated structures following exocytosis. Despite the identification of many molecular components of clathrin-mediated endocytosis, a structural understanding of how these molecules come together to build and retrieve material from the plasma membrane is incomplete. In this aim we sought to directly determine the structure of clathrin-coated vesicles responsible for endocytosis. By understanding how proteins that have been functionally implicated in endocytosis assemble together at the nanoscale we place decades of biochemistry, cell biology, and genetics into a physical model of membrane retrieval. First, using our high-throughput TIRF imaging pipeline discussed above we mapped how over 80 proteins associated with endocytic clathrin-coated structures in PC12 cells. This provided us with the fundamental protein signature of clathrin structure across the plasma membrane. From this analysis we determined that a core group of 30 protein associate with clathrin-coated sites in these cells. This information provided us with a molecular framework to understand the regulation of endocytosis in these cells. Next, we developed a new super-resolution correlative light and electron microscopy imaging method. This development allows us to image the nanometer-scale location of proteins in the context of their local cellular environment. Specifically, we succeeded in developing a robust pipeline for imaging the plasma membrane of cells with both 3D iPALM (interferometric photoactivation localization microcopy) and 3D transmission electron microscopy (TEM) of platinum replicas. In these studies we have imaged the 3D position of the endocytic protein Epsin at single clathrin-coated structures in PC12 cells. From this work we discovered that Epsin sits at the equator of endocytic vesicles where it likely aids in the retrieval of clathrin-coated pits. We have continued to map dozens of other critical endocytic components at the nanoscale with this method to develop a global map of the proteins involved in endocytosis. These studies are allowing us to build structural models for how proteins are organized at single organelles to regulate endocytosis, a central process for all living cells. Aim 3 My lab's major focus is to understand how cellular process work at the molecular scale. To accomplish this, current tools were inadequate. Thus, we focused on the development and application of new imaging methods to uncover the structure of molecules and organelles inside cells. In my group we continue to pioneer four novel imaging technologies. These include, 1) high-throughput microscopy and image processing to map protein distributions across populations of cells, 2) 3D super-resolution fluorescence imaging methods, 3) super-resolution correlative light and electron microscopy to map proteins at the nanoscale, and 4) high-throughput metal ion FRET methods to map distances within individual proteins at the atomic scale. The combination of these methods provides our group with the unique tools to gain a multi-scale understanding of the structure, dynamics, and functions of proteins and protein complexes inside cells.